wild bonobos host geographically restricted malaria parasites...

ARTICLE

Wild bonobos host geographically restrictedmalaria parasites including a putative new LaveraniaspeciesWeimin Liu1, Scott Sherrill-Mix1,2, Gerald H. Learn1, Erik J. Scully3,4, Yingying Li1, Alexa N. Avitto1,

Dorothy E. Loy1,2, Abigail P. Lauder2, Sesh A. Sundararaman1,2, Lindsey J. Plenderleith5, Jean-Bosco N. Ndjango6,

Alexander V. Georgiev3,7, Steve Ahuka-Mundeke8, Martine Peeters9, Paco Bertolani10, Jef Dupain11,

Cintia Garai12, John A. Hart12, Terese B. Hart12, George M. Shaw1,2, Paul M. Sharp 5 & Beatrice H. Hahn1,2

Malaria parasites, though widespread among wild chimpanzees and gorillas, have not been

detected in bonobos. Here, we show that wild-living bonobos are endemically Plasmodium

infected in the eastern-most part of their range. Testing 1556 faecal samples from 11 field

sites, we identify high prevalence Laverania infections in the Tshuapa-Lomami-Lualaba (TL2)

area, but not at other locations across the Congo. TL2 bonobos harbour P. gaboni, formerly

only found in chimpanzees, as well as a potential new species, Plasmodium lomamiensis sp.

nov. Rare co-infections with non-Laverania parasites were also observed. Phylogenetic rela-

tionships among Laverania species are consistent with co-divergence with their gorilla,

chimpanzee and bonobo hosts, suggesting a timescale for their evolution. The absence of

Plasmodium from most field sites could not be explained by parasite seasonality, nor by

bonobo population structure, diet or gut microbiota. Thus, the geographic restriction of

bonobo Plasmodium reflects still unidentified factors that likely influence parasite

transmission.

DOI: 10.1038/s41467-017-01798-5 OPEN

1 Department of Medicine, University of Pennsylvania, Philadelphia, PA 19104, USA. 2Department of Microbiology, University of Pennsylvania, Philadelphia,PA 19104, USA. 3 Department of Human Evolutionary Biology, Harvard University, Cambridge, MA 02138, USA. 4Department of Immunology and InfectiousDiseases, Harvard T.H. Chan School of Public Health, Boston, MA 02115, USA. 5 Institute of Evolutionary Biology and Centre for Immunity, Infection andEvolution, University of Edinburgh, Edinburgh EH9 3FL, UK. 6 Department of Ecology and Management of Plant and Animal Resources, Faculty of Sciences,University of Kisangani, BP 2012 Kisangani, Democratic Republic of the Congo. 7 School of Biological Sciences, Bangor University, Bangor LL57 2UW, UK.8 Institut National de Recherche Biomedicale, University of Kinshasa, BP 1197 Kinshasa, Democratic Republic of the Congo. 9 Unité Mixte Internationale 233,Institut de Recherche pour le Développement (IRD), INSERM U1175, University of Montpellier 1, BP 5045, Montpellier 34394, France. 10 Leverhulme Centrefor Human Evolutionary Studies, University of Cambridge, Cambridge CB2 1QH, UK. 11 African Wildlife Foundation Conservation Centre, P.O. Box 310, 00502Nairobi, Kenya. 12 Lukuru Wildlife Research Foundation, Tshuapa-Lomami-Lualaba Project, BP 2012 Kinshasa, Democratic Republic of the Congo. Weimin Liuand Scott Sherrill-Mix contributed equally to this work. Correspondence and requests for materials should be addressed toB.H.H. (email: [email protected])

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African apes are highly endangered, requiring noninvasiveapproaches to study infectious agents in wild-livingcommunities. To elucidate the origins and evolution of

human malaria parasites, we1–3 and others4–6 have developedPCR-based methods that permit the faecal-based detection andmolecular characterisation of related parasites in wild-living apes.Such studies have shown that chimpanzees (Pan troglodytes) andwestern gorillas (Gorilla gorilla) harbour a plethora of Plasmo-dium parasites, which fall into two major groups7. One group(subgenus Plasmodium) includes several Plasmodium speciesinfecting monkeys as well as ape parasites that are closely relatedto human P. malariae, P. ovale and P. vivax7. Of these, ape P.vivax is known to infect both chimpanzees and gorillas, whilecontemporary human P. vivax represents a lineage that emergedfrom these parasites as it spread out of Africa2. The other group(subgenus Laverania) includes ape parasites that are most closelyrelated to human P. falciparum7. There are currently six describedape Laverania species, which appear to exhibit strict host speci-ficity in wild ape populations1,3–5. These include P. reichenowi(also termed C1), P. gaboni (C2), and P. billcollinsi (C3), whichinfect chimpanzees, as well as P. praefalciparum (G1), P. adleri(G2), and P. blacklocki (G3), which infect western gorillas. Ofthese, only the gorilla parasite P. praefalciparum has crossed thespecies barrier to humans, resulting in the emergence of P. fal-ciparum1,7,8. Although initially based primarily on mitochondrialsequences1, this taxonomy of Laverania species has subsequentlybeen confirmed by analysis of multiple nuclear gene sequences3,9.

Laverania infections have been documented at multiple loca-tions throughout the ranges of chimpanzees and western lowlandgorillas (G. g. gorilla), with estimated prevalence rates in infected

communities ranging between 22 and 40%7. Similarly, apeP. vivax is found in all chimpanzee subspecies as well as westernand eastern gorillas (G. beringei), although estimated prevalencerates are lower, ranging between 4 and 8%7. Studies of Asianprimates have shown that the distribution and prevalence ofPlasmodium infections depends on a number of ecological vari-ables, such as forest cover10, population density11, vector capa-city12 and environmental conditions13, many of which areinterrelated. Although the factors that promote and sustainmalaria transmission in wild apes remain largely unknown, it isclear that Plasmodium species are not uniformly distributedamong them. For example, eastern gorillas harbour ape P. vivax,but do not seem to carry Laverania parasites1,2. More strikingly,bonobos (Pan paniscus) appear to be free of all known apePlasmodium species, despite the screening of multiplecommunities1,2.

The seeming absence of Plasmodium infections from wildbonobos has remained a mystery. Anopheles vectors, includingforest species such as A. moucheti, A. marshallii and A. vinckei,which are known to carry ape Plasmodium parasites14,15 appearto be distributed throughout the bonobo range16. Bonobos arealso very closely related to chimpanzees, suggesting a similarsusceptibility to Plasmodium infection. Finally, there is no evi-dence that bonobos are inherently resistant to Plasmodiumparasites, since human P. falciparum and P. malariae have beendetected in the blood of several captive individuals17. Reasoningthat previous studies may have missed infected communities, weconducted a more extensive survey, increasing both the numberand geographic diversity of sampled bonobo populations. Here,we show that wild bonobos are, in fact, susceptible to a wide

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Fig. 1 Plasmodium infections of wild-living bonobos. Ape study sites are shown in relation to the ranges of the bonobo (P. paniscus, dashed red) and theeastern chimpanzee (P. t. schweinfurthii, dashed blue), with white dots indicating sites where no Plasmodium infection was found (see Table 1 andSupplementary Table 3 for a list of all field sites and their code designation). The Tshuapa–Lomami–Lualaba (TL2) site where bonobos are endemicallyinfected with multiple Plasmodium species, including a newly discovered Laverania species (B1), is shown in red with two dots indicating sampling on bothsides of the Lomami River. Eastern chimpanzee field sites with endemic P. reichenowi, P. gaboni and/or P. billcollinsi infections are shown in yellow. A redcircle highlights one bonobo (KR) and one chimpanzee (PA) field site where B1 parasite sequences were detected in a single faecal sample. Forested areasare shown in dark green, while arid or semiarid areas are depicted in brown. Major lakes and rivers are shown in blue. Dashed yellow lines indicate nationalboundaries. The scale bar indicates 200 km

ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/s41467-017-01798-5

2 NATURE COMMUNICATIONS | 8: 1635 |DOI: 10.1038/s41467-017-01798-5 |www.nature.com/naturecommunications


variety of Plasmodium parasites, including a previously unknownLaverania species that appears specific to bonobos. However,endemic infection was only detected in the eastern-most part ofthe bonobo range, indicating that most wild-living communitieshave lost these parasites.

ResultsBonobos are naturally Laverania infected. Bonobos are found inthe rain forests of the Congo Basin in the Democratic Republic ofthe Congo (DRC). Separated from eastern chimpanzees (P. t.schweinfurthii) and eastern lowland gorillas (G. b. graueri) by theCongo River, their range extends from the Lualaba River in theeast, to the Kasai and Sankuru Rivers in the south, and the LakeTumba and Lake Mai-Ndombe regions in the west (Fig. 1). Initialstudies failed to identify Plasmodium infections in wild bonobos,but were conducted at only two locations (LK and KR)1.Although subsequent surveys included additional bonobo fieldsites (ML, LA, IK, BN, BJ, TL2), faecal samples were only testedfor P. vivax-like parasites2. Here, we screened these (n= 646) aswell as newly collected (n= 803) faecal samples from the same(LA, IK) and additional (LG, BX, MZ) study sites for Laveraniainfection (Fig. 1). Using conventional (diagnostic) PCR to amplifya 956 bp mitochondrial cytochrome B (cytB) fragment1, we failedto detect parasite sequences in 1418 samples from 10 of these 11locations (Table 1). Surprisingly, however, 16 of 138 faecal spe-cimens from the Tshuapa-Lomami-Lualaba (TL2) project sitewere Laverania positive as determined by direct ampliconsequencing (Table 1).Reasoning that conventional PCR screening may have

missed low-level Laverania infection, we retested all availablecytB-negative faecal specimens by subjecting them to anintensified PCR protocol. Since most ape faecal samples containlimited quantities of parasite DNA, we reasoned that testingmultiple aliquots of the same DNA preparation would increasethe likelihood of parasite detection. To avoid PCR contamination,only initially negative samples were re-tested using the intensifiedapproach. Performing 8 to 10 independent PCR reactions foreach DNA sample, we identified 17 additional faecal samplesfrom TL2 to contain cytB sequences, resulting in a total of

33 positive specimens from 24 different apes (Table 1).Although in most cases only one or a few replicates yielded anamplification product (Supplementary Table 1), the intensifiedPCR approach more than doubled the number of positives at theTL2 site, revealing an overall Laverania prevalence of 38%(Table 1). However, this was not observed for other bonobo fieldsites. Intensified PCR of the remaining 1105 samples identifiedonly a single additional positive specimen from the KokoloporiReserve (KR). Thus, malaria parasites are either absent or belowthe limits of faecal detection at the vast majority of bonobo fieldsites.

A new bonobo-specific Laverania species. Having identifiedLaverania-positive bonobo samples, we next sought to molecu-larly characterise the infecting parasites. Since apes are frequentlyco-infected with multiple Plasmodium species, we used limitingdilution PCR, also called single genome amplification (SGA), togenerate mitochondrial cytB sequences (956 bp) devoid of Taqpolymerase-induced artefacts such as in vitro recombination18.Using this approach, we generated 166 limiting dilution-derivedcytB sequences from 34 Laverania-positive bonobo samples,including a unique haplotype from the single positive KR speci-men (Supplementary Table 2). Phylogenetic analysis showed thatthese bonobo parasites fell into two well-supported clades withinthe Laverania subgenus (Fig. 2). One of these comprised a sub-lineage of P. gaboni (C2E) previously found in eastern chim-panzees (P. t. schweinfurthii) in the DRC3. Within this sublineage,bonobo and chimpanzee parasite sequences were completelyinterspersed, indicating that P. gaboni productively infects both ofthese Pan species (Fig. 2 and Supplementary Fig. 1). The otherclade represented a distinct Laverania lineage (B1) that includedonly bonobo parasites, except for a single cytB sequence pre-viously identified3 in an eastern chimpanzee sample (PApts368)east of the Congo/Lualaba River (Fig. 1).To determine whether B1 parasites were more widespread

among eastern chimpanzees than previously recognised, weused regular and intensified PCR to screen faecal samples(n= 562) from nine study sites located closest to the bonoborange (Fig. 1). Although this analysis yielded twice as many

Table 1 Noninvasive screening of wild-living bonobo communities for Laverania infections

Field sitesa Conventional cytB screen Intensified cytB screenb

Samplestestedc

Samplespositived

Detection rate(%)

Samplestestedc

Samplespositived

Detection rate(%)

Combined detectionrate (%)

Balanga (BN) 84 0 0 84 0 0 0Bananjale (BX) 1 0 0 1 0 0 0Bayandjo (BJ) 2 0 0 2 0 0 0Ikela (IK) 465 0 0 432e 0 0 0Kokolopori (KR) 69 0 0 69 1 1.4 1.4f

Lingunda-Boyela (LG) 25 0 0 25 0 0 0Lomako (LA) 307 0 0 289e 0 0 0Lui-kotal (LK) 38 0 0 38 0 0 0Malebo (ML) 262 0 0 n/a n/a n/a n/aManzana (MZ) 165 0 0 165 0 0 0Tshuapa–Lomami–Lualaba(TL2)

138 16 11.6 122 17 13.9 23.9f

n/a not availableaField sites are designated by a two- or three-letter code (their location is shown in Fig. 1). Regular cytB screening data for KR and LK sites have previously been reported2bSamples initially negative by conventional (diagnostic) cytB PCR were subsequently retested by intensified PCR, performing 8–10 additional amplification reactions per faecal DNA (see SupplementaryTable 1 for details)cThe host species origin of all faecal samples was confirmed by mitochondrial DNA (D-loop) analysis. As previously reported, the number of individuals tested at the KR, LK and TL2 sites was 38, 17 and63, respectively3dAll amplification products were sequence confirmed to represent Laverania parasiteseOnly a subset of the originally screened IK and LA samples were available for intensified PCRfOf a total of 38 and 63 bonobos at the KR and TL2 sites1, 2, 1 and 24 were Laverania infected, indicating a prevalence of 2.6% and 38%, respectively

NATURE COMMUNICATIONS | DOI: 10.1038/s41467-017-01798-5 ARTICLE




Laverania positive samples as conventional PCR (SupplementaryTable 3), none of the newly derived cytB sequences fell withinthe B1 clade (Supplementary Table 4). Instead, eastern chimpan-zees were exclusively infected with P. reichenowi (C1), P. gaboni(C2) and P. billcollinsi (C3) (Supplementary Fig. 1). These

data indicate that TL2 bonobos harbour a form of P. gabonithat is highly prevalent in neighbouring eastern chimpanzeesas well as a second Laverania species that seems unique tobonobos.To characterise the newly identified bonobo parasites in other

regions of their genomes, we used SGA to target additionalorganelle and nuclear loci for analysis (Supplementary Table 2).These included 3.4 and 3.3 kb mitochondrial DNA (mtDNA)fragments, which together span the entire mitochondrial genome;a 390 bp caseinolytic protease M (clpM) gene fragment from theapicoplast genome; and three nuclear loci, including portions ofgenes encoding the erythrocyte binding antigens 165 (eba165;790 bp) and 175 (eba175; 394 bp), and the gametocyte surfaceprotein P47 (p47; 800 bp). Phylogenetic analyses of 134 newlyderived parasite sequences yielded very similar results (withrespect to the clustering of parasites into major clades) in allgenomic regions (Fig. 3 and Supplementary Fig. 2). Except for asingle C1 eba175 sequence indicative of a rare P. reichenowiinfection (Supplementary Fig. 2d), all other bonobo-derivedsequences fell either within P. gaboni or the B1 clade(Supplementary Table 2). This new clade was supported by highbootstrap values in all genomic regions analysed, except for theshort eba175 fragment. It also consistently grouped as a sisterclade to P. reichenowi. These findings, along with the extent ofgenetic divergence between P. reichenowi and the newly identifiedbonobo parasite clade, argue strongly for the existence of anadditional Laverania species that is specific for bonobos (Figs. 2and 3 and Supplementary Figs. 1 and 2). The finding of B1 cytB(Fig. 2) and eba165 (Fig. 3a) parasite sequences in a singlechimpanzee faecal sample collected 280 km east of TL2 doesnot argue against this, since it shows that B1 parasites reachedthis geographic region, but failed to spread in the residentchimpanzee population (Supplementary Table 4). We propose toname the new bonobo parasite species Plasmodium lomamiensissp. nov. to highlight its discovery in Lomami National Park,using faecal-derived mitochondrial, apicoplast and nuclearparasite sequences as the type material (Supplementary Data 1)19. Although classifying ape Laverania species solely on thebasis of genetic information has been controversial3,17,20, thereare no obvious alternatives given the endangered status of wildapes, the prevalence of mixed Laverania species infections(Supplementary Table 2) and the cryptic nature of theseparasites3,9,21.

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Fig. 2 Relationship of bonobo parasites to ape Laverania species.A maximum likelihood tree of mitochondrial cytochrome B (cytB)sequences (956 bp) depicting the phylogenetic position of newly derivedbonobo parasite sequences (magenta) is shown. Only distinct cytBhaplotypes are depicted (the full set of SGA-derived bonobo parasitesequences is shown in Supplementary Fig. 1). Sequences are colour-coded,with capital letters indicating their field site of origin (see Fig. 1 for locationof field sites) and lowercase letters denoting their host species andsubspecies origin (ptt: P. t. troglodytes, red; pte: P. t. ellioti, orange; pts:P. t. schweinfurthii, blue; ggg: G. g. gorilla, green; pp: Pan paniscus, magenta).C1, C2 and C3 represent the chimpanzee parasites P. reichenowi, P. gaboniand P. billcollinsi; G1, G2 and G3 represent the gorilla parasites P.praefalciparum, P. adleri and P. blacklocki (the P. falciparum 3D7 referencesequence is shown in black). P. reichenowi (C1) and P. gaboni (C2)mitochondrial sequences are known to segregate into two geographicallydefined subclades according to their collection site in 'western' (W) or'eastern' (E) Africa3. Bonobo parasite sequences (magenta) cluster withP. gaboni from eastern chimpanzees (C2E), but also form a new clade,termed B1. The tree was constructed using PhyML58 with TIM2+I+G as theevolutionary model. Bootstrap values are shown for major nodes only (thescale bar represents 0.01 substitutions per site)




TL2 bonobos also harbour non-Laverania parasites. SGA ofbonobo faecal DNA also yielded rare sequences from non-Laverania parasites that resulted from primer cross-reactivity(Supplementary Table 2). One such cytB sequence clustered witha previously characterised parasite sequence from a chimpanzeesample (DGptt540), forming a well-supported lineage that wasonly distantly related to human and ape P. malariae (Fig. 4a),while two other clpM sequences clustered with ape and human P.vivax parasites (Fig. 4b). To search for additional non-Laveraniainfections, we used P. vivax- and P. malariae-specific primers torescreen bonobo faecal samples from the BX (n= 1), KR (n= 69),LA (n= 199) and TL2 (n= 138) field sites using intensified PCR.This analysis confirmed P. vivax infection in one bonobo sample,and identified P. vivax and P. ovale curtisi sequences in twoadditional samples, all from the TL2 site (Fig. 4c). Furthercharacterisation revealed that the P. ovale curtisi-positive sample

also contained ape P. vivax sequences (Fig. 4d). Thus, of the 24Laverania-positive bonobos at the TL2 site, 3 also harbouredP. malariae-, P. vivax- and/or P. ovale-related parasites, while anadditional bonobo exhibited a P. vivax monoinfection (Supple-mentary Table 2). Although the recovered sequences were tooshort to differentiate human- and ape-specific parasite lineages,the results show that bonobos, like chimpanzees and gorillas, arefrequently infected with multiple Laverania and non-Laveraniaspecies1,5,7. However, unlike chimpanzees and gorillas, bonobosharbour these parasites in only one particular part of their range.

The Lomami River is not a barrier to malaria transmission.Analysing mtDNA sequences to determine the populationstructure of wild bonobo populations, two previous studiesreported that the Lomami River, but not other tributaries of the

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Fig. 3 A new Laverania species specific for bonobos. a, b Maximum likelihood phylogenetic trees are shown for nuclear gene fragments of the a erythrocyte-binding antigen 165 (eba165; 790 bp) and b the gametocyte surface protein P47 (p47; 800 bp) of Laverania parasites. Sequences are labelled and coloured asin Fig. 2 (identical sequences from different samples are shown; identical sequences from the same sample are excluded). C1, C2 and C3 represent thechimpanzee parasites P. reichenowi, P. gaboni and P. billcollinsi; G1, G2 and G3 represent the gorilla parasites P. praefalciparum, P. adleri and P. blacklocki(PrCDC and Pf3D7 reference sequences are shown in black). Bonobo parasite sequences cluster within P. gaboni (C2) or form a new distinct clade (B1),indicating a new Laverania species (see text for information on the single eba165 B1 sequence from an eastern chimpanzee). The trees were constructedusing PhyML58 with TPM3uf+G (a) and GTR+G (b) as evolutionary models. Bootstrap values are shown for major nodes only (the scale bar represents0.01 substitutions per site)





Congo River, represents a geographical barrier to bonobo geneflow22,23. We thus considered the possibility that bonobos in thewestern and central regions of the DRC had acquired a malariaprotective trait that had not spread to bonobo populations east ofthe Lomami River. To investigate this, we subjected Plasmodium-positive and -negative samples from TL2 to the same hostmtDNA analysis (Supplementary Data 2) and compared theresulting haplotypes to all previously reported bonobo mtDNAsequences (Fig. 5a and Supplementary Fig. 3). Phylogeneticanalysis showed that most of the newly derived mtDNAsequences from TL2 (blue) fell into two clades that were exclu-sively comprised of sequences from bonobos sampled east of theLomami River (Supplementary Fig. 3b)22,23. However, 4 new TL2haplotypes representing 15 faecal samples, including 4 Laverania-positive specimens, did not fall within these two 'eastern' clades(arrows in Fig. 5a and Supplementary Fig. 3a). Analysis of theirGPS coordinates revealed that they were all collected west (TL2-W) of the Lomami River (Fig. 5b). These results thus confirm andextend previous findings showing that bonobos east of theLomami River represent a genetically (at least matrilineally)

isolated population22,23. However, this isolation does not explainthe geographic restriction of bonobo malaria, since Laverania-positive individuals were found on both sides of the LomamiRiver. Although it remains unknown how far the Plasmodiumendemic area extends beyond TL2 in the eastern Congo, it seemsclear that the Lomami River itself does not represent a barrier tomalaria transmission.

Climate does not explain the distribution of bonobo malaria.Because climatic factors such as ambient temperature andrainfall are known to influence malaria transmissionin humans24–26, we asked whether seasonal differences inPlasmodium prevalence could explain the lack of parasite detec-tion at the majority of bonobo study sites. Comparison ofsample dates across all field sites revealed no obvious associationbetween faecal parasite positivity and the month of specimencollection (Table 2). For example, samples collected inNovember and December at the TL2 site included a large fractionof malaria-positive specimens, but this was not the case forsamples collected during these same months at the IK, KR and LA

P. malariae

P. vivax

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MSggg7183

EKggg514

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TL2pp3853

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cox1

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DSggg83

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DPptt143DSggg83

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EKggg1179

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NKggg1167

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P. vivax

P. hylobati P. inui

Fig. 4 Bonobo infections with non-Laverania parasites. Maximum likelihood phylogenetic trees are shown for mitochondrial and apicoplast gene sequencesof non-Laverania parasites. Ape-derived a cytB (956 bp), b clpM (327 bp), c cox1 (296 bp) and d clpM (574 bp) sequences are labelled and coloured as inFig. 2 (identical sequences from different samples are shown; identical sequences from the same sample are excluded). Human and monkey parasitereference sequences from the database are labelled by black squares and circles, respectively. Brackets indicate non-Laverania species, including P.malariae, P. vivax, P. ovale curtisi and P. ovale wallikeri (available sequences are too short to differentiate ape- and human-specific lineages) as well as themonkey parasites P. inui and P. hylobati. Newly identified bonobo parasite sequences are indicated by arrows, all of which are from the TL2 site. One TL2cytB sequence clusters with a previously reported parasite sequence from a chimpanzee sample (DGptt540), forming a well-supported lineage that is onlydistantly related to human and ape P. malariae, and thus likely represents a new P. malariae-related species. The trees were constructed using PhyML58 withGTR+G (a), TRN+I (b, d) and TIM2+I (c) as evolutionary models. Bootstrap values over 70% are shown for major nodes only (the scale bar represents0.01 substitutions per site)




field sites. To examine the impact of climatic variation on bonoboparasite detection more directly, we used a statistical modelshown to be strongly predictive of spatiotemporal variation inLaverania infection among wild-living chimpanzees (Erik Scully,unpublished results). This model, which was parameterised usingPCR screening data from 2436 chimpanzee faecal samples col-lected at 55 locations across equatorial Africa7, showed thatambient temperature, daily temperature fluctuations and forestcover, but not rainfall, each influenced the probability of Laver-ania detection.

Using only specimens with known sampling dates and GPScoordinates for which land surface temperature and forest coverdata were also available (Supplementary Table 5), we estimatedthe probability of Laverania infection for each of the 11 bonobofield sites. Assuming similar climatic influences on chimpanzeeand bonobo parasite development and transmission, this analysisshowed that at seven sites for which a sufficiently large numberof samples were available, bonobos were significantly less

frequently Laverania infected than predicted by the climatemodel. For the BN, IK, LA, LK and MZ sites, the model predicteda less than one in a million probability that a positive samplewould not be detected if bonobos at these sites exhibited similarinfection patterns as chimpanzees. Moreover, for the KR site,where only one sample was Laverania positive, seasonal variationcould not explain this very low detection rate (Table 2). Therate of parasite detection at the TL2 site, where 27 of 113 sampleswith climate data were positive, was lower than, but notsignificantly different from, that predicted for a chimpanzeestudy site with similar ecological conditions. The very smallsample sizes at BJ and BX sites lacked statistical power to detectdifferences, and the low predicted probability of infection atthe ML site indicated that more sampling during months ofhigher infection probabilities would be necessary to confidentlyreject the climate model. Nevertheless, our sampling density atmost sites was sufficient to conclude that the scarcity of infectionin bonobos was not caused by biased sampling during seasonal

mtDNA D-loop

a blyondji

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Fig. 5 The Lomami River is not a barrier to Laverania parasite transmission. a Maximum likelihood phylogenetic tree of bonobo mitochondrial (D-loop)sequences. Haplotypes are labelled by field site (see Fig. 1 and refs. 7,22,23,77, for their geographic location and code designation), with those identified atmultiple field sites indicated (e.g., C/Wamba/KR/BN/IK/LA). Newly derived haplotypes from the TL2 site are shown in blue (previously reported mtDNAsequences are shown in black)22,23,77. Brackets highlight two clades that are exclusively comprised of mtDNA sequences from bonobos sampled east ofthe Lomami River. TL2 haplotypes that do not fall within these clades (denoted by arrows) were all sampled west of the Lomami River (TL2-W). The treewas constructed using PhyML58 with HKY+G as the evolutionary model. Bayesian posterior probability values ≥ 0.6 are shown (the scale bar represents0.01 substitutions per site). b Locations of individual bonobo faecal samples collected at the TL2 site. Sampling locations west (TL2-W) and east (TL2-Eand TL2-NE) of the Lomami River were plotted using GPS coordinates, with red and white dots indicating Laverania parasite positive and negativespecimens, respectively. Samples that contained P. reichenowi, P. malariae-like, P. vivax-like and P. ovale-like parasites are also indicated. Forested areas areshown in green, while savannas are depicted in brown. The Lomami River is shown in blue. Local villages are denoted by black squares. The scale barindicates 2 km





troughs (Table 2). Thus, it appears that seasonal or climaticvariation in parasite prevalence can be excluded as an explanationfor the observed geographic restriction of bonobo Plasmodiuminfections.

Bonobo diet is not associated with faecal parasite detection.Wild apes consume a variety of plants, fruits, barks and piths,some of which have been reported to have antimalarialactivity27–29. We thus asked whether our inability to detectPlasmodium infections at most bonobo field sites was due to thepresence of certain plants, which upon ingestion would reduceparasite titres below the limits of faecal detection. To examine thispossibility, we selected a subset of Laverania-positive (n= 18) and-negative (n= 51) bonobo faecal samples from endemic (TL2)and non-endemic (KR, IK, LG, LK) field sites, and characterisedtheir plant content by targeting two regions of the chloroplastgenome for high-throughput sequencing (SupplementaryTable 6). These comprised a 500 bp fragment of the rbcL gene anda 750 bp fragment of the matK gene, both of which have beenused as barcodes to identify land plants30–32, including in stoolsamples from endangered species33. Laverania-positive (n= 14)and -negative (n= 15) chimpanzee faecal samples were analysedfor control (Supplementary Table 6).

Samples were sequenced to a mean depth of 16,054 matK and21,995 rbcL paired-end reads, which were clustered intooperational taxonomic units (OTUs) and assigned to taxonomicgroups using a custom matK and rbcL reference database(Supplementary Fig. 4). Using a permutational multivariateanalysis of variance (PERMANOVA) to compare unweightedUniFrac distances34 as a measure of large-scale differences inplant composition, we found small differences between faecalsamples from bonobos and chimpanzees (matK: 2.0% of variance,p= 0.003; rbcL: 2.8% of variance, p< 10−6), but substantialdifferences between faecal samples from different study sites(matK: 19.8% of variance, p< 10−6; rbcL: 18.6% of variance, p<10−6). However, no significant differences were observed betweenLaverania-positive and -negative faecal samples (matK: 1.2% ofvariance, p= 0.18; rbcL: 0.8% of variance, p= 0.71), suggestingthat the lack of parasite detection was not associated with theabundance of certain plant phyla in the diet (Fig. 6a, b andSupplementary Fig. 5).

We also compiled a list of 466 African plant species(Supplementary Table 7), which have been reported to havepotential antimalarial activity29,35,36, and looked for related matKand rbcL sequences in bonobo faecal samples from endemic andnonendemic field sites. Although a BLAST search identified 65matK and 490 rbcL OTUs that shared 95% sequence identity with3 and 17 of these putative antimalarial species, respectively, nonewas significantly more abundant at field sites where Laveraniainfections were absent (Supplementary Fig. 6). Similar resultswere obtained when the remaining plant OTUs were comparedbetween endemic and nonendemic bonobo field sites. Finally, nocompositional differences were observed in the plant content ofLaverania-positive and -negative chimpanzee faecal samples(Fig. 6a, b). Although these analyses provide only a snapshot ofbonobo and chimpanzee plant diet, they failed to identify anassociation between particular plant constituents and parasitedetection in faecal samples.

The faecal microbiome does not predict Laverania infection.Plasmodium infections have been reported to influence thebacterial communities in the gut, with certain parasitescausing intestinal dysbiosis37 and certain gut microbiota enhan-cing the host’s anti-parasite immune responses38. To examinepotential interactions between the faecal microbiome andLaverania infection in bonobos, we used the same samplesselected for plant analyses (Supplementary Table 6) for bacterial16S rRNA sequencing (Laverania-positive and -negativechimpanzee samples again served as a control). Samples weresequenced to a mean depth of 65,132 reads, which were clusteredinto OTUs and assigned to taxonomic groups (SupplementaryFig. 7a). Examining Shannon diversity as a marker ofdysbiotic outgrowth or loss of bacterial taxa, we failed to findsignificant differences in within-sample (alpha) diversity betweenspecimens from Laverania-positive and -negative bonobos(or chimpanzees), or between specimens from endemic (TL2) andnonendemic (KR, IK, LG, LK) field sites (Supplementary Fig. 8a).Using unweighted UniFrac distance to compare between-sample(beta) diversity34, we found that as previously reported39 bonoboand chimpanzee faecal microbiomes differed in their bacterialcomposition (Fig. 6c and Supplementary Figs. 7b and 8b). Sam-ples from the same field site were also often compositionally moresimilar to each other than to samples from other field

Table 2 Seasonal variation in parasite prevalence does not explain the geographic restriction of bonobo malaria

Site Sample collectiona LaveraniacytB

Model predictions

Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec Pos Total Meanb Significance cutoffc p-valued

BN 49 35 0 84 28.2 17 <10–6

BX 1 0 1 0.4 0 1.0BJ 2 0 2 0.7 0 1.0IK 107 22 46 4 85 7 61 121 0 453 147 121 <10–6

KR 5 4 9 (1) 13 33 1 64 25.1 15 <10−6

LG 11 14 0 25 6.9 2 0.003LA 12 43 30 73 41 0 199 54.5 39 <10−6

LK 4 6 28 0 38 14.2 7 <10−6

ML 56 86 0 142 4.1 0 0.176MZ 5 49 9 16 57 25 0 161 22.6 12 <10−6

TL2 8 (4) 8 64 (18) 33 (5) 27 113 39.5 27 0.083

aOnly samples with known collection dates and GPS coordinates, for which land surface temperature and forest cover data could also be obtained, were included in the climate analysis (sample numbersare thus lower than in Table 1). Numbers in brackets indicate Laverania cytB-positive samplesbThe expected mean number of positive samples predicted by the climate modelcThe significance cutoff corresponds to the 0.45% (5%, with Bonferroni correction for 11 tests) of the Poisson binomial distribution for the probabilities predicted by the climate model. Values of 0indicate that a greater sample size is necessary to confidently reject the climate modeldA significant Bonferroni adjusted p-value (p< 0.05) indicates that climate can be rejected as the explanation for rare or absent Plasmodium infections at any particular sampling site




sites (Supplementary Fig. 8b and c). Using PERMANOVA toexamine the sources of this variation, we found that ape speciesaccounted for 7.4% (p< 10−6), study site for 19.3% (p< 10−6)and Laverania positivity for 1.2% of the variance (p= 0.043),respectively. Considering only chimpanzee samples, studysite accounted for 17.8% (p= 0.000018) and Laverania positivityfor 4.0% of variance (p= 0.25). Comparing only samplesfrom TL2 bonobos, differences among three sample locations(Fig. 5b) accounted for 14.6% (p< 10−6) and Laverania infectionfor 4.5% of variance (p= 0.0023). Thus, there was a small butsignificant compositional difference between the faecal micro-biome of Laverania- positive and -negative bonobos at TL2(the lack of significance in chimpanzees may be due to a smallersample size).Using Wilcoxon rank sum tests to look for OTUs that

were driving these differences, we found one assigned to thefamily Ruminococcaceae that was significantly depleted, and twoothers assigned to family Lachnospiraceae and Prevotella coprithat were significantly enriched in Laverania-positive TL2bonobo samples (Supplementary Fig. 9). However, comparingsamples from TL2 to nonendemic sites did not yield significantlyhigher UniFrac distance values than comparing samples betweenthese nonendemic sites (Supplementary Fig. 8b). Thus, while theabundance of some bacterial taxa differed slightly betweenLaverania-positive and -negative bonobos at TL2, compositionaldifferences between samples from TL2 and nonendemic siteswere no greater than expected between any two random sites,thus failing to provide a microbial signature of Laveraniainfection for that site.

DiscussionA complete account of Plasmodium infections in wild Africanapes, including their host associations, prevalence, geographicdistribution and vector preferences, is critical for understandingthe origins of human malaria and gauging future zoonoticrisks. Previous studies documented numerous Plasmodiumspecies in wild chimpanzees and gorillas, but failed to find similarinfections in wild bonobos1,2. Here, we show that bonobos har-bour a multitude of Plasmodium species, although endemicinfection is limited to only a small part of their range east of theLomami River. Analyses of climate data and parasite seasonality,as well as host characteristics, including bonobo populationstructure, plant consumption and faecal microbiome composi-tion, failed to provide an explanation for this geographic

restriction. Thus, other factors must be responsible for the unevendistribution of bonobo Plasmodium infections, including thepossibility of a protective mutation that has not spread east of theLomami River.Studies in Asia have shown that both species richness and

prevalence of primate malarias are closely linked to the habitat offorest-dwelling Anopheles, rather than the distribution of theprimates themselves10,12. Thus, factors that negatively impact thebreeding conditions, development and distribution of transmit-ting vectors may be responsible, at least in part, for the absence ofPlasmodium infections at most bonobo sites. Ecological factorsmay also influence bonobo density or other behaviours that affectvector exposure. For example, captive orangutans, which live inhigher group densities than their wild counterparts, also havehigher rates of Plasmodium infection11. Finally, it is conceivablethat bonobos at the Plasmodium-negative sites carry otherinfections that induce cross-protective immunity or compete forthe same resources40. Bonobos are clearly susceptible to a varietyof Plasmodium species. Thus, examining why neither Laveranianor non-Laverania infections are sustained throughout much ofthe bonobo range may identify new drivers of vector dynamics orother transmission risks that could aid malaria eradication effortsin humans.Although diagnostic PCR of matched blood and stool samples

from Plasmodium-infected captive macaques indicated faecaldetection rates of up to 96%41, parasite detection in wild primatesis much less sensitive due to widely varying sample quality. Whilethe new bonobo Laverania infections were identified by con-ventional PCR (Table 1), we reasoned that multiple PCR repli-cates would increase the chances of parasite detection and thusthe number of sequences for phylogenetic analyses. This wasindeed the case since 18 (of 191) bonobo and 46 (of 517)chimpanzee samples identified as Plasmodium negative by con-ventional PCR were subsequently found parasite positive byintensified PCR (Table 1 and Supplementary Table 3). Usingthese results (Supplementary Table 1), we estimated the sensi-tivity of a single PCR replicate to be 16.7% (95% confidenceinterval (CI): 12.9–20.8), and the sensitivity of 8 and 10 replicatesto be 76.8% and 83.9%, respectively (see Methods). Althoughlabour intensive, costly and prone to contamination, intensifiedPCR is the method of choice for samples with low parasite levels,such as partially degraded specimens from remote field sites, sinceits increased sensitivity can detect rare, and possibly even new,Plasmodium species.

matKa rbcLb 16S rRNA

−0.2 −0.1 0.0 0.1 0.2 0.3

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e)Fig. 6 Laverania infection of bonobos is not associated with particular faecal plant or microbiome constituents. A principal component analysis ofunweighted UniFrac distances was used to visualise compositional differences of a, b plant (matK and rbcL) and c bacterial (16S rRNA) constituents inLaverania-positive (dark border) and -negative (light border) faecal samples from bonobos (blue) and chimpanzees (pink). The sample positions (shownfor the first two components) do not indicate separate clustering of Laverania-positive and -negative samples





Phylogenetic analyses of over 3000 mitochondrial, apicoplastand nuclear parasite sequences from chimpanzee andgorilla faecal samples have consistently pointed to the existenceof six ape Laverania species1,3–6. Here, we propose a seventhspecies, P. lomamiensis, on the basis of partial organelle andnuclear gene sequences derived from faecal DNA. Unlike otherape parasites, this species seems to be bonobo specific andgeographically restricted to the Lomami River basin. Although atraditional full taxonomic description might seem preferable,the highly endangered status of wild bonobos precludes bloodcollection. Moreover, the cryptic nature of Laverania parasitesas well as the high frequency of mixed species infections renders amorphological description uninformative. Indeed, P. gaboniand P. reichenowi, which are morphologically indistinguishablefrom P. falciparum, were only recently shown by full-lengthgenome sequencing to represent distinct, non-recombiningspecies9,42, and the same type of analysis uncovered thathuman P. ovale comprises not one but two sympatric parasitespecies43,44. Thus, a true description of cryptic Plasmodiumspecies must come from multilocus genetic analysis. Although weused single template amplification to derive high-quality orga-nelle and nuclear sequences of multiple P. lomamiensis strains,the number of loci and samples is still limited. As DNA-onlyspecies taxonomy is gaining wider acceptance19, it will be inter-esting to determine how much genetic information is necessary toreliably classify P. lomamiensis and other putative Plasmodiumspecies.The newly identified bonobo parasites prompt speculation

about the causes and timescale of Laverania diversification. Whenthe only Laverania species characterised were P. falciparum andP. reichenowi, it was widely assumed that these two species hadco-diverged with their hosts45,46, placing their common ancestorat the same time as the common ancestor of humans andchimpanzees around 6–7Mya47. This hypothesis was under-mined by the finding of additional Laverania species, in particularthe discovery that P. falciparum resulted from the recent hostswitch of the gorilla parasite P. praefalciparum1. However, P.praefalciparum and P. reichenowi could have co-diverged with theancestors of gorillas and chimpanzees. The phylogenetic position(Figs. 2 and 3) of the newly described bonobo parasite, P.lomamiensis (B1), which is more closely related to P. reichenowi(C1) than to P. praefalciparum (G1), provides a triad of parasitespecies with the same relationships as their hosts (SupplementaryFig. 10). It is thus tempting to speculate that this clade arosethrough host–parasite co-divergence. Under this scenario, thecommon ancestor of P. reichenowi and P. praefalciparum wouldhave existed approximately 8–9Mya47, an estimate that is 2 to 4times older than some have concluded from molecular clockanalyses for the equivalent divergence of P. reichenowi and P.falciparum48,49. P. reichenowi and P. lomamiensis would havediverged around 2Mya47. Molecular clocks for Laverania speciesmay not be very precise: for example, P. reichenowi and P.praefalciparum are clearly not 4 times more divergent than P.reichenowi and P. lomamiensis (Figs. 2 and 3). However, giventhat P. reichenowi and P. gaboni are about 3 times more divergentthan P. reichenowi and P. praefalciparum9, it is possible that thecommon ancestor of the entire Laverania clade existed around25–30 Mya.The co-divergence scenario also predicts that the ancestor of

today’s bonobos was infected with the ancestor of P. lomamiensis,which was subsequently lost from most bonobo populations. TheCongo River, which forms the boundary between the ranges ofchimpanzees and bonobos (Fig. 1), is thought to have existedsince long before the divergences among African apes50; yet,somehow the ancestor of bonobos reached the left bank of theCongo. This may have happened during one of several periods of

aridity when river levels might have been low enough to permitthe crossing in the northeast of the current bonobo range50.Furthermore, mitochondrial DNA analyses suggest that theremay have been an early population split between the ancestors ofbonobos now found east and west of the Lomami River51. Thus,the loss of P. lomamiensis from western populations may haveoccurred early in bonobo history. It should be noted that theinfection status of bonobos at sites other than TL2 east of theLomami (e.g., BJ and BX; Fig. 1) remains unknown, because toofew samples have been collected. However, bonobos immediatelywest of the Lomami at TL2 must have reacquired P. lomamiensis,indicating that the river is not a barrier to mosquitoes from theeast, and that western bonobos as a whole do not share agenetically based resistance to infection. The co-divergence sce-nario also implies that a related parasite was lost from the humanlineage, which might have been due to an early human populationbottleneck, an ancestral hunter–gatherer lifestyle52 and/or the lossof the gene that synthesises N-glycolylneuraminic acid (Neu5Gc),which may have affected the ability of the parasite to infecthuman erythrocytes53.

The extent of divergence (Figs. 2 and 3) between P. gaboni (C2)and P. adleri (G2) is similar to that between P. reichenowi and P.praefalciparum, suggesting that the former pair may also have co-diverged with their hosts. Within the C2/G2 clade there is againno human parasite species, and the only bonobo parasites in thislineage clearly reflect recent transmissions of P. gaboni fromeastern chimpanzees, rather than co-divergence. In this case, theloss of a putative B2 lineage from bonobos is not surprising giventhe loss of P. lomamiensis from most of the bonobo range. Thelack of close relatives of P. billcollinsi (C3) and P. blacklocki (G3)would also be indicative of past losses of parasite lineages fromparticular ape hosts. Although the processes that contributed tothe emergence of today’s Laverania lineages remain unknown, itseems likely that both co-divergence and cross-species transmis-sion events shaped their evolutionary history, as has beenobserved for many other pathogens.One characteristic of Laverania parasites infecting wild apes is

their highly specific host tropism. This species specificity is notshared by non-Laverania parasites, such as P. vivax, which infectsbonobos, humans, chimpanzees and gorillas2,54. However, evenwithin the Laverania subgenus, host specificity is not absolute.Bonobos at TL2 are commonly infected with the chimpanzeeparasite P. gaboni, which appears to have crossed the LualabaRiver on multiple occasions (Fig. 2). Bonobos are also susceptibleto the chimpanzee parasite P. reichenowi (Supplementary Fig. 2d),while eastern chimpanzees appear susceptible to the bonoboparasite P. lomamiensis (Figs. 2 and 3), although both of thesecross-infections appear to reflect rare events that fail to result inonward transmission. Thus, on the one hand, Laverania speciesare extremely host specific, implying strong barriers to cross-species transmission, and on the other hand there is evidence thaton occasion these barriers can be overcome. Given the very closegenetic relationship of chimpanzees and bonobos, examples ofcross-infections are perhaps not surprising. However, the findingthat in captive settings bonobos can become infected with humanP. falciparum17, while chimpanzees can harbour gorilla parasitesand vice versa55, indicates that Laverania host specificity goesbeyond incompatibilities of receptor–ligand interactions duringerythrocyte invasion56. While ape Laverania parasites have notyet been detected in humans8,57, it seems clear that themechanisms governing host specificity are complex and thatsome barriers are more readily surmountable than others. Giventhe new bonobo data, it will be critical to determine exactly howP. praefalciparum was able to jump the species barrier to humans,in order to determine what might enable one of the other apeLaverania parasites to do the same.




MethodsApe samples. Faecal samples from wild-living bonobos and eastern chimpanzeeswere obtained from existing specimen banks, or were newly collected at previouslyreported1–3 as well as new study sites (LG, BX, MZ) in the DRC (Fig. 1). While allavailable bonobo samples (n= 1556) were analysed, eastern chimpanzee specimens(n= 580) were selected from 9 field sites most proximal to the bonobo range.All samples were obtained noninvasively from apes in remote forest areas, pre-served (1:1 vol/vol) in RNAlater, transported at ambient temperatures and stored at−80 °C. Faecal DNA was extracted using the QIAamp Stool DNA mini kit (Qiagen)and all specimens were subjected to host mtDNA analysis to confirm their speciesand subspecies origin1–3. The latter analysis also gave an indication of samplequality, which confirmed that samples from Laverania-negative field sites werenot any more degraded than samples from TL2. For the KR, LK and TL2 field sites,the number of sampled individuals has previously been determined by micro-satellite analyses2. All samples were obtained with approval from the Ministries ofScientific Research and Technology, the Department of Ecology and Managementof Plant and Animal Resources of the University of Kisangani, the Ministries ofHealth and Environment, and the National Ethics Committee in the DRC, andshipped in compliance with Convention on International Trade in EndangeredSpecies of Wild Fauna and Flora regulations and country-specific import andexport permits.

Conventional and intensified PCR. Bonobo and chimpanzee faecal samples werefirst screened for Laverania parasites by conventional (diagnostic) PCR, targeting a956 bp mitochondrial cytB fragment using primers DW2 (5′-TAATGCCTAGACGTATTCCTGATTATCCAG-3′) and DW4 (5′-TGTTTGCTTGGGAGCTGTAATCATAATGTG-3′) in the first round, and Pfcytb1 (5′-CTCTATTAATTTAGTTAAAGCACA-3′) and PLAS2a (5′-GTGGTAATTGACATCCWATCC-3′) in the secondround of PCR as previously described1. Since this approach tests only a singlealiquot of each faecal DNA, we reasoned that parasites present in low concentra-tions may have been missed. To increase the sensitivity of parasite detection, wethus tested 8 to 10 aliquots of the same DNA preparation using the same primersand amplification conditions. To guard against false positives, only samples thatwere negative by conventional PCR were subjected to the intensified PCRscreening. Assuming that PCR amplification was 100% specific, that all PCRreactions were independent with a fixed probability of detecting a positive sample,and that samples found negative in all PCR replicates could still be Plasmodiumpositive, we expected that the number of positive PCR reactions for a faecal samplewould follow a zero-truncated binomial distribution and that the likelihood of thedata would be the product over all samples with a positive reaction for thisintensified PCR approach. Therefore, we used numerical optimisation to find themaximum likelihood estimate of sensitivity for these data and determined 95% CIsusing likelihood ratios. Using intensified PCR data from both bonobo and chim-panzee samples (Supplementary Table 1), we estimated the sensitivity of a singlePCR replicate to be 16.7% (95% CI: 12.9–20.8), and thus the sensitivity of 8 and 10independent PCR reactions to be 76.8% and 83.9%, respectively. Separate analysesof chimpanzee and bonobo samples yielded similar sensitivity estimates for singlePCR replicates: 17.5% (95% CI: 13.0–22.5) for eastern chimpanzees and 14.6%(95% CI: 8.47–22.3) for bonobos. Intensified PCR was also used to screen 199bonobo faecal samples from the TL2, KR and LA field sites for non-Laveraniainfections using parasite-specific primer sets. P. vivax primers targeted a 296 bpcox1 fragment using Pv2768p (5′-GTATGGATCGAATCTTACTTATTC-3′)and Pv3287n (5′-AATACCAGATACTAAAAGACCAACAATGATA-3′) in thefirst round, and Pv2856p (5′-CTTATTACAAATTGCAATCATAAAACTTTAGGT-3′) and Pv3185n (5′-TCCTCCAAATTCTGCTGCTG TAGATAAAATG-3′) in the second round of PCR as previously described8. P. malariae specificprimers targeted a 600 bp mitochondrial cytB fragment using Pm4659p (5′-ATTTATTATCTTCAATTCCAGCACTT-3′) and Pm5501n (5′-GCATGTTAACTCGATAAATACTAA-3′) in the first round, and Pm4740p (5′-ATTACATTTTATACTTCCATTTGTTGC-3′) and Pm5369n (5′-TTCAGAAATATCGTCTTATCGTAGC-3′) in the second round of PCR. P. vivax-specific primers detected bothP. vivax and P. ovale, while P. malariae-specific primers amplified only positivecontrol samples (one P. malariae-positive sample was detected due to the cross-reactivity of regular cytB primers; Fig. 4a). All amplicons were sequenced directlywithout interim cloning.

Single genome amplification. To derive Plasmodium sequences devoid of PCR-induced errors, all PCR-positive bonobo and chimpanzee faecal samples weresubjected to SGA as previously described1,2,18. According to the Poisson dis-tribution, the DNA dilution that yields PCR products in no more than 30% of wellscontains one amplifiable template per positive reaction more than 80% of the time.Faecal DNA was thus end point diluted in 96-well plates, and the dilution thatyielded <30%-positive wells was used to generate single template-derivedsequences. For Laverania-positive bonobo samples, mitochondrial (cytB; 3.4 and3.3 kb mitochondrial half genomes), apicoplast (clpM) and nuclear (eba165, eba175and p47) gene regions were amplified using previously reported primer sets andamplification conditions (Supplementary Table 2)1–3,8,56. For Laverania-positivechimpanzee samples, only the 956 bp mitochondrial cytB fragment was amplified(Supplementary Table 4). One bonobo faecal sample (TL2.3874) positive for P.

ovale curtisi by conventional PCR also yielded a P. vivax-specific 574 bp apicoplastclpM fragment when subjected to SGA analysis.

Phylogenetic analyses. Sequences were aligned using CLUSTAL W (version 2.1),visually inspected and regions that could not be unambiguously aligned wereremoved from subsequent analyses. Maximum likelihood phylogenetic trees andbootstrap support were estimated using PhyML (version 3.0)58, which infersevolutionary model parameters and phylograms concurrently. Evolutionary modelswere selected using jModelTest (version 2.1.4)59. Bayesian posterior probabilitieswere determined using MrBayes (version 3.2.4)60 using two simultaneous inde-pendent analyses with a 25% burn-in. Convergence was determined when theaverage deviation of split frequencies was <0.01.

Climate model of ape Laverania infection. To evaluate whether the absence orlow prevalence of Laverania infection at most bonobo sampling sites could beexplained by seasonal variation in parasite transmission, we developed a logisticgeneralised linear mixed model to infer the probability of parasite detection foreach sample relative to the climatic variables observed at the time of specimencollection. Briefly, this model, which incorporates mean ambient temperature (AT),daily temperature variation (TV) and percent forest cover (FC), was parameterisedusing 2436 chimpanzee faecal samples from 55 sampling sites across equatorialAfrica7 and found to be strongly predictive of Laverania infection in wild chim-panzees (Erik Scully, unpublished results). Assuming similar relationships betweenclimatic variables and infection probability in chimpanzees and bonobos, andincluding only samples for which climate data were available, we inferred thepredicted probability of Laverania infection for each bonobo sample using theequation:

Predicted Probability ¼ 1

1þ e� interceptþ0:355 ´AT�0:164 ´AT2þ0:032 ´ FCþ0:208 ´TVð Þ

where intercept is −2.538 for samples screened using conventional PCR (i.e., onereplicate) and −1.374 for those screened using intensive PCR (i.e., 8–10 replicates),and AT, TV and FC are each corrected by subtracting the means of the chimpanzeedata set (23.4 for AT, 9.1 for TV and 77.2 for FC). For each bonobo sample, weused MODerate Resolution Imaging Spectroradiometer (MODIS) and daytime andnight-time land surface temperature (LST) data sets61,62 in one-day temporalresolution (MOD11A1) after applying the minimum/maximum air temperaturetransformations as previously described24 to derive (1) the mean ambient airtemperature and (2) the mean daily air temperature fluctuation. Each of thetemperature variables was calculated as the average of LST measurements takenduring the period 30 days prior to sample collection. Forest cover data wereextracted from high-resolution global maps as previously described63. For eachsampling site, the mean and ranges of these ecological variables are summarised inSupplementary Table 5. Assuming that each specimen is independent and has aprobability of detected infection as assigned by the climate model, the number ofpositives observed at a given site will be a sum of Bernoulli variables with varyingprobabilities and thus should follow the Poisson binomial distribution64. We cal-culated the cumulative probability of seeing less than or equal the observed numberof positive samples64 given the set of climate estimates for each site to generate p-values and used Bonferroni correction to account for multiple comparisons. A lowp-value indicates that climatic variation is very unlikely to account for the observedscarcity of infection.

Characterisation of faecal plant composition. Chloroplast ribulose bisphosphatecarboxylase large chain (rbcL) and maturase K (matK) gene regions are widelyused as bar codes for land plants30–32 and were thus selected to characteriseplant components in Laverania-positive and negative bonobo (n= 78) and chim-panzee (n= 20) faecal samples (Supplementary Table 6). Faecal DNA was extractedusing the PowerSoil-htp 96 Well Soil DNA Isolation Kit (MO BIO Laboratories,Carlsbad, CA, USA). We modified rbcL primers previously reported to have highplant discriminatory ability32 for MiSeq sequencing by adding an Illumina adapter(underlined). These included rbcLbF (5′-AGACCTWTTTGAAGAAGGTTCWGT-3′) and rbcLbR (5′-TCGGTYAGAGCRGGCATRTGCCA-3′) for the firstround of PCR, and R1_rbcL634F (5′-TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGATGCGTTGGAGAGACCGTTTC-3′) and R2_rbcLbR (5′-GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGTCGGTYAGAGCRGGCATRTGCCA-3′) for the second round of PCR. We also modified matK primers recentlyimproved to achieve high PCR success rates31 in a similar fashion, using matK390F(5′-CGATCTATTCATTCAATATTTC-3′) and matK1326R (5′-TCTAGCACACGAAAGTCGAAGT-3′) in the first round of PCR, and R1_matK472F (5′-TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGCCCRTYCATCTGGAAATCTTGGTTC-3′) and R2_matK1248R (5′-GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGGCTRTRATAATGAGAAA GATTTCTGC-3′) in the second round ofPCR.

Amplification for both rbcL and matK gene regions was performed using 2.5 μlof sample DNA in a 25 μl reaction volume containing 0.5 μl dNTPs (10 mM of eachdNTP), 10 pmol of each first round primer, 2.5 μl PCR buffer, 0.1 μl BSA solution(50 μg/ml), and 0.25 μl Expand Long Template enzyme mix (Expand LongTemplate PCR System) for the first round of PCR. Cycling conditions included





an initial denature step of 2 min at 94 °C, followed by 15 cycles of denaturation(94 °C, 10 s), annealing (45 °C, 30 s) and elongation (68 °C, 1 min), followed by25 cycles of denaturation (94 °C, 10 s), annealing (48 °C, 30 s) and elongation(68 °C, 1 min; with 10 s increments for each successive cycle), followed by a finalelongation step of 10 min at 68 °C. For the second round PCR, 2 μl of the firstround product was used in 25 μl reaction volume. Cycling conditions includedan initial denature step of 2 min at 94 °C, followed by 25 cycles of denaturation(94 °C, 10 s), annealing (52 °C, 30 s) and elongation (68 °C, 1 min), followed by afinal elongation step of 10 min at 68 °C. For each faecal sample, rbcL and matKgene regions were amplified in duplicate, the products were pooled, purifiedusing QIAquick Gel Extraction Kit and sequenced using the Illumina Miseq v2(500 cycle).

Sequence reads were separated by barcode, quality filtered for an expectednumber of errors <1 and an exact match to primer sequences, and the 5′ and 3′reads of each pair were concatenated after trimming off primer sequences. OTUswere formed using Swarm65, and OTUs containing only a single read discarded.Representative sequences of each OTU were aligned using MAFFT66 and aphylogenetic tree was inferred using FastTree67. To create a database for taxonomicassignment, all reads matching the search terms 'matK' or 'rbcL' were downloadedfrom the European Nucleotide Archive and indexed in a BLAST database. Thisdatabase was searched using a representative sequence from each OTU, andtaxonomy was assigned as the most specific taxonomic rank shared by all BLASThits with a total bit score within 98% of the best hit. Samples with <5000 reads wereremoved from the analysis.

Characterisation of faecal bacterial constituents. The same faecal DNAsamples used for plant analyses were also subjected to bacterial 16S rRNA genesequencing (Supplementary Table 6). The 16S rRNA gene amplification wasperformed as previously described68, using 5 μl of faecal DNA, the AccuPrimeTaq DNA Polymerase High Fidelity System (Thermo Fisher), and V1V2 regionprimers containing Illumina adapters, barcode and linker regions. Each faecalsample was amplified in four independent reactions, with the products pooledand purified using AMPure XP beads (Beckman Coulter) before sequencingusing Illumina MiSeq v2 (500 cycle). Sequences were separated by barcode,and paired reads were merged using bbmerge69. Reads were clustered intoOTUs using a cutoff of 97% identity and taxonomically assigned using QIIMEv1.9.1 and the Greengenes database70,71. OTUs formed from single reads werediscarded. Samples with <15,000 sequences per sample were removed from theanalysis.

Statistical analyses. All analyses were performed in R v3.3.372. Within-sample(alpha) diversity was calculated using the Shannon diversity index73. Between-sample (beta) diversity of matK and rbcL data was calculated using unweightedUniFrac distances after rarefaction to 5000 reads per sample34. Between-samplediversity of 16S rRNA data was also calculated using unweighted UniFrac, but afterrarefaction to 15,000 reads per sample34. We opted to use unweighted distancevalues because they permit the examination of rare taxa that might be related to thephenotype examined; however, weighted UniFrac as well as weighted andunweighted Bray–Curtis dissimilarity values gave comparable results (matk: allMantel tests r > 0.34, p < 10−6; rbcl: all Mantel tests r > 0.68, p < 10−6; 16S: allMantel tests r> 0.77, p < 10−6). Unweighted UniFrac distances were also used forprincipal coordinates analysis74, t-distributed stochastic neighbour embedding75

and permutational analysis of variance (http://CRAN.R-project.org/package=vegan)76. Analyses of 16S rRNA data revealed that TL2 bonobo samplesformed three distinct clusters (Supplementary Fig. 8c), corresponding to threedifferent sampling locations west (TL2-W) and east (TL2-E and TL2-NE) of theLomami River (Fig. 5b). To control for site-specific differences in faecal plantcomposition, we measured depletion of matK and rbcL OTUs between samplesfrom TL2-E, TL2-NE and TL2-W and three non-endemic LK, KR and IK field sitesusing Wilcoxon rank sum tests. The p-values from the nine pairwise comparisonswere combined using Fisher’s method with the test statistic and degrees of freedomdivided by 3 to control for correlation between tests. Changes in bacterial OTUproportions between TL2 Laverania-positive and -negative samples were measuredusing Wilcoxon rank sum tests.

Nomenclatural acts. This published work and the nomenclatural acts it containshave been registered in ZooBank, the proposed online registration system for theInternational Code of Zoological Nomenclature (ICZN). The ZooBank LSIDs (LifeScience Identifiers) can be resolved and the associated information viewed throughany standard web browser by appending the LSID to the prefix 'http://zoobank.org/'. The LSID for this publication is: urn:lsid:zoobank.org:act:BF143A7B-DA74-469D-B44C-40B5EF82B2DB.

Code availability. Analysis code is archived on Zenodo (https://zenodo.org/record/886174) at DOI: (http://doi.org/10.5281/zenodo.886174).

Data availability. Newly derived Laverania and non-Laverania parasite sequencesas well as bonobo mtDNA haplotypes have been deposited in GenBank underaccession numbers KY790455-KY790593 (also see Supplementary Data 1 and 2).

High-throughput plant and microbiome sequences are archived in the NCBISequence Read Archive (SRA) under BioProject PRJNA389566.

Received: 5 June 2017 Accepted: 16 October 2017

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AcknowledgementsWe thank the staff of the TL2 site, the Institut National de Recherches Biomédicales(INRB, Kinshasa, DRC) and the Bonobo Conservation Initiative for field work in theDRC; Richard Carter for helpful discussions; the Ministry of Scientific Research andTechnology, the Department of Ecology and Management of Plant and AnimalResources of the University of Kisangani, the Ministries of Health and Environment andthe National Ethics Committee for permission to collect samples in the DRC. This workwas supported by grants from the National Institutes of Health (R01 AI 091595, R01 AI058715, R01 AI 120810, R37 AI 050529, T32 AI 007532, T32 AI 007632, P30 AI 045008),the Agence Nationale de Recherche sur le Sida (ANRS 12125/12182/12255), the AgenceNationale de Recherche (Programme Blanc, Sciences de la Vie, de la Santé et des Eco-systémes and ANR 11 BSV3 021 01, Projet PRIMAL), Harvard University and the ArthurL. Greene Fund.

Author contributionsAll authors contributed to the acquisition, analysis and interpretation of the data; W.L.,S.S.-M., G.M.S., P.M.S. and B.H.H. conceived, planned and executed the study; J.-B.N.N.,A.V.G., S.A.-M., M.P., P.B., J.D., C.G., J.A.H. and T.B.H. conducted or supervisedfieldwork; W.L., Y.L., D.E.L., A.N.A. and S.A.S. performed noninvasive ape Plasmodiumtesting; W.L. and A.P.L. performed faecal plant and microbiome analyses; E.J.S. devel-oped a climate model predictive of ape Laverania infection; S.S.-M. performed statisticaland bioinformatic analyses; G.H.L., L.J.P. and P.M.S. performed phylogenetic analyses;



http://modis.gsfc.nasa.gov/

http://modis.gsfc.nasa.gov/

http://jgi.doe.gov/data-and-tools/bbtools/bb-tools-user-guide/bbmerge-guide/

http://jgi.doe.gov/data-and-tools/bbtools/bb-tools-user-guide/bbmerge-guide/

http://www.R-project.org/

http://www.R-project.org/



W.L., S.S.-M., G.M.S., P.M.S. and B.H.H. coordinated the contributions of all authors andwrote the manuscript.

Additional informationSupplementary Information accompanies this paper at doi:10.1038/s41467-017-01798-5.

Competing interests: The authors declare no competing financial interests.

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